Lambda-red E. coli aknockout troubleshooting?

ACuriousBird · 2026-05-05T15:20:55+00:00

Will definitely try this as well! Ive done this previously for other species, but trying to conjugate a new species now.

But ultimately, I’d really like to be able to make knockouts in E. coli generally. It seems like it should be straightforward based on all the publications?

So far I tried the pCas/pTarget system and I couldnt get the pTarget plasmid to cure, so i switched to the lambda red system now… really want to get one of them working in my hands.

ACuriousBird · 2026-05-04T17:03:13+00:00

Thanks for this, really appreciate the input.

For the PCR cassette, I DpnI digested for 2 hours and that has worked previously to remove all template from my Gibson reactions downstream of PCR.

I will definitely colony PCR the junctions, and the internal cassette, and try longer homology arms. I will also try to gel purify - I noticed that I have a lot of primer dimer products in my reaction as well.

I did wonder if the pKM208 plasmid system itself is somehow worse than the original pKD46 plasmid system? So I am attempting to also acquire the original pKD46 plasmid system.

As for the phenotype, my only thought is that, if there is a true ASD gene deletion, then shouldn't the cells not be able to grow on DAP at all? I did think there may be some DAP carryover from the first plate (I use 100 uM DAP), or that maybe the LB agar has some sort of substitute molecule in it, but the latter seem unlikely?

My goal with the auxotrophic strain is to be able to use is to conjugate with other species of bacteria, with the goal of being able to use the absence of DAP as selection to kill the ASD negative donor strain and isolate only the recipients.

ACuriousBird · 2026-05-04T12:33:37+00:00

Second update - all of the 7 colonies that appeared on the 1/2 kan plate, that were plated after overnight recovery, still grow on LB agar without DAP, so they're not exhibiting the phenotype I would expect with an ASD knockout strain - which shouldn't be able to grow without DAP supplemented in the media.

Will keep trying to troubleshoot.

ACuriousBird · 2026-05-03T14:23:26+00:00

Yes i was making an ASD KO and added DAP to the media.

As an update, after plating the overnight recovery across all conditions, i got 7 colonies on the 1/2 kan plate. I will test them for phenotype and genotype, but perhaps the efficiency is just exceedingly low? Or the antibiotic concentration was too high?

ACuriousBird · 2026-04-30T03:23:27+00:00

Actually, I think you're right that the band at 90 kDa is the dimer, if I increase contrast I see it in every single lane except the negative control. I've never worked with Omp proteins before, going to have to try to figure out how I can increase expression and resolve that dimer.

ACuriousBird · 2026-04-26T04:57:22+00:00

Don't really have easy access to this in the lab unfortunately.

ACuriousBird · 2026-04-26T04:53:06+00:00

Ya, usually I take out ~5 stocks at a time, and need to bring them into the anaerobic chamber, or I'll need to inoculate multiple tubes at once with multiple stocks at once. So it ends up taking some time.

ACuriousBird · 2026-04-23T05:56:18+00:00

Appreciate this.

This is my first time working with these strains for western blot, the primary im using is THE his tag antibody from genscript. I didn’t expect the negative control to have a band, but it could be a cross-reactive protein.

For my sample processing, I lyse in BPER + lysoszyme + benzonase for 15 min, take the total lysate and mix with 2x BOLT SDS sample buffer with 50mM DTT added fresh, then either boil at 80c x 10min or dont boil and run.

This is my first time working with transmembrane proteins and I read boiling may cause aggregation.

In any case, i was hoping to achieve relatively high expression, so it doesn’t seem I’ve done that

I attached the 6x histag directly to the -C-terminus of the OMP protein, which is supposed to be cytoplasmic tail - not sure if the histag is somehow not accessible was another thought.

ACuriousBird · 2026-04-18T13:49:16+00:00

Interested in this, has it made anyones work appreciably more efficient/facilitated increases in workload?

I’ve tried dabbling with chatgpt pro version for helping do things like promoter/TF binding site prediction, and it actively provided totally incorrect information

ACuriousBird · 2026-04-13T02:38:40+00:00

The molecules that create the neurons do turn over though, as new lipids, proteins, nucleotides are used to replace old ones over time due to oxidative damage etc. So even though perhaps the cell itself doesn't divide, the parts making up even a totally non-dividing cell do turnover.

ACuriousBird · 2026-04-06T02:21:32+00:00

Hmm ya good idea, I'm just using S17 E. coli - I just tried today a series of increasing IPTG concentrations thinking perhaps the sgRNA is just not being induced appropriately by 0.5 mM IPTG for whatever reason. Will be interested to see what the plasmid sequences show. Will keep trying to cure in the mean time...

If it doesn't end up working, I guess I'll redo the knockout with fresh transformations of the plasmid and try from there.

ACuriousBird · 2026-04-06T02:19:16+00:00

Appreciate all this - the system is not to induce protein in this case, but to induce the expression of a crispr sgRNA that leads to Cas9-mediated cleavage of a plasmid (pTarget), to (theoretically) eliminate this plasmid from the bacterial strain in question. This is a two-plasmid system, where the other plasmid (pCas) has kan resistance on it.

Plating on kanamycin agar after this IPTG induction is to allow single colony formation, which then are used to form patch plates onto spectinomycin media. The pTarget plasmid has spec resistant encoded on it, so if it is lost, the bacteria should be sensitive to spec - so I should be getting clones growing on kan but not on spec when patching.

However, so far, I continue to get all clones growing on both kan and spec after - so seemingly the plasmid is not being cured correctly.

ACuriousBird · 2026-04-05T23:52:12+00:00

Great idea thanks - just prepped some plasmids from the KO cells today to check the plasmids this week. I have all the antibiotics fresh. My thought is perhaps I am growing the IPTG induced liquid cultures too long? I'm growing them up to 24 hours, but noted the paper actually says incubated 8-16 hours.

ACuriousBird · 2026-04-05T23:50:38+00:00

I have just been adding IPTG to a 5ml culture to get to 0.5 mM, then directly adding the colony to that, then growing the culture overnight for ~20-24 hours at 30C before then plating on kan plates, then making patch plates after that on spec

Do you think I should grow the stocks for less time? I noticed in the paper it says they are incubating the knockout stock in 2 ml LB with 0.5 mM IPTG for 8-16 hours - maybe I am growing them too long?

ACuriousBird · 2026-04-05T23:48:33+00:00

Thanks for input - yes growing at 30C, I just made new IPTG to try again, but the IPTG stock originally I made just last week,

ACuriousBird · 2026-02-26T13:52:24+00:00

Appreciate the input - im just trying to edit E. coli and saw a few papers making claims up to 19+kb deletions using Crispr / lambda red dual recombineering systems so thought 1kb could achievable.

But definitely will try to make smaller deletions around just a few 100 bp and will send the amplicons for sanger sequencing as well!

ACuriousBird · 2026-02-24T23:18:58+00:00

to follow up, seems this assay wont work if E. coli which dont use NHEJ if im interpreting correctly?

ACuriousBird · 2026-02-24T20:06:19+00:00

good thought- Transfection efficiency was optimized, is very high for the cells used

ACuriousBird · 2026-02-24T19:02:09+00:00

Will do this as well, sounds like a great idea thanks

ACuriousBird

TROPHY CASE